Abstract

Perturbation in the Dystroglycan (Dg)–Dystrophin (Dys) complex results in muscular dystrophies and brain abnormalities in human. Here we report that Drosophila is an excellent genetically tractable model to study muscular dystrophies and neuronal abnormalities caused by defects in this complex. Using a fluorescence polarization assay, we show a high conservation in Dg–Dys interaction between human and Drosophila. Genetic and RNAi‐induced perturbations of Dg and Dys in Drosophila cause cell polarity and muscular dystrophy phenotypes: decreased mobility, age‐dependent muscle degeneration and defective photoreceptor path‐finding. Dg and Dys are required in targeting glial cells and neurons for correct neuronal migration. Importantly, we now report that Dg interacts with insulin receptor and Nck/Dock SH2/SH3‐adaptor molecule in photoreceptor path‐finding. This is the first demonstration of a genetic interaction between Dg and InR.

Mutations in Dystrophin glycoprotein complex (DGC) in vertebrates lead to muscle degeneration as well as phenotypes in many other cell types (Durbeej and Campbell, 2002; Cohn, 2005). For example, several muscular dystrophies exhibit neuronal migration disorders (Muntoni et al, 2002; Qu and Smith, 2004), showing that Dg interactions are essential for normal neuron migration. However, the mechanism of action and regulation of this complex are not fully understood in any cell type. Multiple proteins interacting with Dg have been identified through biochemical assays resulting in the hypothesis that Dg is involved in regulation of the actin cytoskeleton, signal transduction and cell morphology (Yang et al, 1995; Sotgia et al, 2001; Spence et al, 2002, 2004a, 2004b).

It is now critical to analyze which of these biochemical interactions are required for Dg–Dys function and regulation and in which cell types do these interactions take place. Model organisms are essential for these functional studies and a few of such models exist and have been analyzed. For example, Dys is defective in Duchenne Muscular Dystrophy (DMD) patients as well as in mdx mice, the highly studied mouse model for DMD. However, in mdx mice, a compensating process limits muscular necrosis during most of the animal's life (Durbeej and Campbell, 2002; Michele et al, 2002; Moore et al, 2002). In addition, Caenorhabditis elegans and zebrafish have recently been used to model muscular dystrophies (Gieseler et al, 2000; Parsons et al, 2002; Bassett and Currie, 2003).

Dys is a 427 kDa rod‐shaped protein that is defective in DMD. The huge gene encodes for three full‐length dystrophin isoforms and four shorter, truncated products, controlled by different internal promoters. The complex structure of the gene is highly conserved during evolution. Similarly to the mammalian dystrophin gene, the fly gene encodes three full‐length dystrophin‐like products (DLPs) and three truncated products consisting of the C‐terminal and cysteine‐rich domains with various extensions into the spectrin‐like repeats domain of DLP. Like the human gene products, the Drosophila gene products are expressed in a tissue‐specific manner (Neuman et al, 2001, 2005; Figure 1A).

Dg and Dys interact in vivo, setting up the anterior–posterior polarity of the oocyte. (A) The Drosophila Dys gene structure. Bars represent exons, horizontal bold lines—introns, bent arrows—translation start sites. Dys genetic deletion 8‐2 and UAS RNAi Dys constructs are shown. UAS dsDys C‐term will affect all known Dys transcripts while UAS dsDys RB is specific for the short transcript RB and UAS dsDys N‐term for the transcripts RA and RC (additional transcripts have recently been isolated (Neuman et al, 2005)). (B) A cartoon of Drosophila Dg–Dys complex is shown. The transmembrane protein Dg provides a link between Laminin in the extracellular matrix and Dys that is attached to the intracellular cytoskeleton. Dg C‐terminal Dys‐binding peptide shown in Figure 2C is marked in red. (C–E) The oocyte polarity marker Orb, which colocalizes with MTOC, is mislocalized in the oocytes of Dg mutants (D, hsFLP; FRT42D Dg323/FRT42D Ubi‐GFP), Dys mutant (Dys8‐2/DysKX43;Table I) and transgenic dsDys flies (E, dsDys N‐term/MatTub‐GAl4;Table I). Instead of being localized to the posterior of the developing oocyte at stages 4–6, Orb surrounds the oocyte in a circle or accumulates in a clump at one side of the oocyte. Red=Orb, Green=DAPI (C, E) or GFP (D). In the right top corners: mislocalization of MTOC in stage 6 oocytes marked by Orb. (F) A bar graph showing that Dg interacts with Dys in the oocyte polarity assay. Transheterozygous Dg323/+; DysKX43/+ and Dg323/+; Dys8‐2/+ mutants show oocyte polarity defects with increased frequency in comparison to control Dg323/+ or Dys KX43/+ flies (Dg323/+ 9.4%, n=117; Dys KX43/+ 5.5%, n=163; Dg323/+; DysKX43/+ 37.4±6.1%, n=309 and Dg323/+; Dys8‐2/+ 37.9%, n=124).

We now report that Drosophila Dg and Dys mutants develop age‐dependent muscle degeneration and mobility defects, indicating that this easy to genetically manipulate organism serves as a remarkably good model for muscular dystrophy. Using this model, we demonstrate that Dg–Dys complex is required in brain in the photoreceptor neurons and in the targeting glial cells for proper axon path‐finding, suggesting that ECM‐based process regulated both from neuronal and glial side contribute to axon migration. Furthermore, the loss‐of‐function‐mutant analysis and genetic interactions suggest that Dg and Dys act in similar axon path‐finding processes as Insulin Receptor (InR) and the adaptor protein Nck/Dock.

Results

Dg and Dys are both required for cellular polarity in Drosophila

A gain of function screen for mutants defective in polarity in Drosophila oogenesis resulted in the finding of Drosophila homologs of components in the Dg complex: Drosophila Dg, LamininA and Dys (Deng and Ruohola‐Baker, 2000; Deng et al, 2003; Figure 1B). Further analysis revealed that the Drosophila genome has all the known components of the Dg complex (Greener and Roberts, 2000; Dekkers et al, 2004). While vertebrates have two closely related proteins, dystrophin and utrophin, encoded by two different genes, Drosophila has only one gene encoding Dystrophin. The expression of Drosophila Dys overlaps with Dg in adult and embryonic tissues (Supplementary Figures 1–3).

To test whether Dg and Dys act in the same cell types and interact genetically in Drosophila, we isolated a mutant allele for Dystrophin and generated dsRNA constructs (Figure 1A). We have analyzed the phenotypes associated with two Dys deletion mutants (Dys8‐2 and Dyse6) and three different dsRNA constructs (Figure 1A; dsDysN‐term targets the long, dsDysRB the short and dsDysC‐term all Dys isoforms) and compared these to the phenotypes of previously isolated Dg mutants (Deng et al, 2003). A significant reduction of Dys was observed with all mutants analyzed (Supplementary Figure 3C–I, M and N; Supplementary Table I). Specifically, while the genetic loss‐of‐function mutant (Dys8‐2/DysKX43) showed a 149‐fold reduction, N‐ and C‐terminal dsDys constructs showed 6–18‐fold reduction of the transcript DLP2 (Supplementary Table I).

Dg is required for cellular polarity: in Dg germline clones, the early oocyte polarity marker Orb fails to show the normal posterior localization in stage 4–6 oocytes (Deng et al, 2003; Table I; Figure 1C and D). This and the accompanied growth defect of the egg chamber can be partially rescued by germ line expression of full‐length Dg protein (Yatsenko et al, 2006). To analyze whether Dys is also required in the germline for oocyte polarity, we examined Orb localization in Dys mutant ovaries (Dys8‐2/DysKX43, Dyse6/DysKX43 or pUASPdsDys N‐term/MatTub‐Gal4). Reduction of Dys function in the germline resulted in an Orb mislocalization phenotype reminiscent of the phenotype seen in Dg mutant; Orb surrounds the entire oocyte in a circle, or it accumulates in a clump at the sides of the oocyte (Figure 1E; Table I). Therefore, Dys, like Dg, is required in germ line for establishment of early oocyte polarity. We also analyzed Dys function in another cell type, follicle epithelial cells, and observed that reduction of Dys results in polarity defects in this cell type as well (Supplementary Figure 3J–L).

To test whether Dg and Dys act in the same process in the germ line, we tested to see if Dg and Dys showed genetic interactions in the oocyte polarity assay; the polarity of Dg323/+; DysKX43/+ and Dg323/+; Dys8‐2/+ oocytes was analyzed. The double heterozygous animals showed significant polarity defects indistinguishable from the homozygous Dg323 or Dys8‐2/DysKX43 mutants, suggesting that Dg and Dys interact in this process (Figure 1F). Thus, both Dg and Dys are required in the germ line and interact in the establishment of cellular polarity during oogenesis.

The Dys–Dg interaction is conserved from human to flies

While the crystal structure of the human Dys–Dg complex has been solved (Huang et al, 2000), the binding affinity for this interaction in human or Drosophila has not been analyzed. We developed a fluorescence polarization assay to determine the binding dissociation constants (Kd) for both the human and Drosophila complexes (Figure 2). In this assay, human or Drosophila Dys (WW+EF hand domains) was titrated into buffer containing fluorescent‐labeled Dg peptide. Human Dys binds human Dg peptide (HmWWbsI) with a Kd of 7.6±1.6 μM. While Drosophila Dys binds Drosophila Dg peptide (DmWWbsI) with a Kd of 16±4 μM (Figure 2A, B and D). To verify that we were measuring binding at the same interface elucidated by the crystal structure, we tested human Dys binding with a mutant Dg peptide in which the tyrosine of the PPPY motif (Tyr 892; Figure 2C) was mutated to a proline (HmWWbsI‐P: KNMTPYRSPPPPVSP). This tyrosine contributes two hydrogen bonds to the binding interface and forms van der Waals contacts to a hydrophobic pocket on the dystrophin WW domain (Huang et al, 2000), all of which are expected to be lost upon mutation to proline. As expected, this titration showed reduced affinity (Kd 172±39 μM) indicating that the assay measures the correct interaction (Figure 2A and D).

Dg and Dys interact in vitro; fluorescence polarization assay reveals that Dys binding to Dg is highly conserved from human to flies. (A) MilliAnisotropy values of human Dys WW+EF hand titrated into buffer containing fluorescently labeled Dg peptides show that human Dys can bind both human (HmWWbsI) and fly (DmWWbsI) Dg peptides. BSA titrated with 200 nM HmWWbsI peptide serves as a negative control. (B) MilliAnisotropy value of Drosophila Dys WW+EF hand titrated into buffer containing fluorescently labeled Dg peptides indicates that Drosophila Dys can also bind both Drosophila (DmWWbsI) and human (HmWWbsI) Dg peptides. BSA is used as the negative control. Binding affinity is reduced in both human and Drosophila models when the wild‐type human peptide is substituted with a mutated peptide (HmWWbsI‐P) in which the terminal tyrosine of the PPxY motif is mutated to a proline. (C) Space filling model of the interaction surface between human Dys WW+EF hand and human Dg peptide HmWWbsI. Human Dys residues that directly contact the HmWWbsI are colored gray. The tyrosine of the PPxY motif, mutated to proline in the HmWWbsI‐P peptide is colored in cyan (arrow). (D) Dissociation constants of human and Drosophila Dys–Dg interaction. Data indicate that this interaction is highly conserved from fly to man: human Dg can interact with Drosophila Dys (WW+EF hand) and vice versa with similar Kd.

In addition, we tested whether human Dg can interact with Drosophila Dys and vice versa. Both these cross species interactions were in the same range as the within species interactions with binding affinities of 24 and 3.7 μM, respectively (Figure 2D). The affinity measured for Dg–Dys interactions is in the expected range for previously analyzed WW‐interactions (Kato et al, 2002). These data show that the Dys–Dg protein interface is highly conserved from humans to flies, suggesting that insights from Drosophila should be transferable to humans.

Dys and Dg mutants show mobility defects

Defects in the Dg complex in human cause muscular dystrophies, which are associated with muscle weakening and degeneration (Cohn and Campbell, 2000). To test whether the Dys–Dg complex plays a similar role in Drosophila muscle function, we first analyzed the mobility of the Drosophila Dg and Dys mutants by measuring their climbing capability (Benzer, 1967) using dsDg and dsDys constructs driven by P‐tub‐Gal4 and the Dys loss‐of‐function mutant Dys8‐2/DysKX43. This rate of climbing decay in Dg and Dys mutants was significantly faster than in wild‐type flies, suggesting that Dg and Dys might be required in the musculature (Figure 3A and B; Table I, T1/2(mobility): control 22–24 days, Dys8‐2/DysKX43 12 days).

Mutations in Dg and Dys cause decreased mobility. (A) Dys and Dg function is required for normal locomotion. Dys and Dg mutants exhibit impaired climbing ability relative to control flies. They begin adult life with normal mobility, but the climbing decay rate is faster in mutants than in wild‐type flies. This indicates that Dg and Dys defects cause age‐dependent climbing disability. (B) A bar‐graph showing that Dys mutant 8‐2 and transgenic ubiquitous Dg and Dys RNAi animals (dsDg and dsDys/P‐tub Gal4) and muscle‐specific dsDg and dsDys/24B‐Gal4 mutants have lost 50% of their climbing ability in 10–14 days after eclosion in comparison to 22–24 days in control.

To test whether the climbing defects in DGC mutant animals were due to Dg and Dys function in muscle tissue, we analyzed mobility of Dys and Dg mutants using dsDg and dsDys constructs driven by mesodermal driver 24B‐Gal4. The speed of climbing decay in dsDg and dsDys/24B Gal4 mutants was similar to what was observed for dsDg and dsDys/P‐tub Gal4 mutants (Figure 3B, T1/2(mobility): control 22–24 days, dsDg/P‐tub‐Gal4 10 days, dsDg/24B‐Gal4 11 days, dsDysC‐term/P‐tub‐Gal4 13 days, dsDysC‐term/24B‐Gal4 10 days). These results indicate that Dg–Dys complex is required in the mesoderm.

Age dependent degeneration of Dys and Dg mutant muscles

To understand the cell biological basis for the observed mobility defects in Dg and Dys mutants, we analyzed their muscle morphology (dsDg and dsDys/P‐tub‐Gal4, Dys8‐2/DysKX43). Histological analysis of the major thoracic muscles showed age‐dependent muscle degeneration in Dg and Dys mutants, consistent with the mobility dysfunction in these mutants (Figure 4A–G; Table I). Confocal and light micrographs of histological sections revealed that the cellular appearance of muscle in Dg and Dys mutants was less organized than in control flies, numerous lesions within the muscular tissue were observed. In 12‐day‐old control flies the indirect flight muscles (IFM) have well structured muscle fibers with peripherally located nuclei (Figure 4A and D). Twelve days after eclosion, animals expressing dsDg and dsDys or Dys8‐2/DysKX43 mutants show loss of muscle fiber organization, vacuolization (Figure 4B, C and G) and absence of some muscles (Figure 4F). These phenotypes became much more pronounced in older mutant flies; the frequency of muscle degeneration increased six‐fold in the mutants (Dys8‐2/DysKX43 or dsDg and dsDys crossed to P‐tub Gal4) compared to the controls during a 9‐day period (Figure 4H; Table I). Similar phenotypes have been observed before in Drosophila parkin and pink1 mutants (Pesah et al, 2004; Yang et al, 2006).

Dg and Dys mutants manifest age‐dependent muscle degeneration. (A–C) Confocal analysis of histological transverse sections of IFM of 12 days old adult flies stained with a nuclear marker DAPI in red. (A) Control flies (UASGFP/P‐tub‐Gal4) show normal organization of the IFM and the muscle fibers are well structured with the nuclei located at the periphery (arrows). (B, C) Dg and Dys mutants (dsDg30A and dsDysN‐term/P‐tub‐Gal4) show severe muscle degeneration: wasting and loss of muscle tissue, vacuolization (asterisks), the integrity of subsets of muscle cells is disrupted and the nuclei appear to be dispersed between fibers (arrows). (D–G, I–K) Light microscopy of histological transverse sections of IFMs stained with H&E. (D) Control (UASGFP/P‐tub‐Gal4, 12 days after eclosion). (E–G) Dys and Dg mutants (dsDys N‐term/P‐tub‐Gal4, dsDg30A/P‐tub‐Gal4) exhibit mainly normal muscle architecture at 3 days after eclosion, but at 12 days in most of the cases the muscle degeneration progresses, the density of myofibrils per muscle decreases and some muscles are absent (arrowhead) or vacuolized (G, asterisk). (H) Bar graph represents increase in frequency of muscle fiber organization defects in 9 days in Dys8‐2and Dg–Dys transgenic animals (dsDg and dsDys/P‐tub‐Gal4) and in 17 days in flies with directed knockout of Dg and Dys in muscle (dsDg and dsDys/24B‐Gal4), which suggest that muscle degeneration has an age‐dependent character. Independent IFMs were calculated (Table I). (I–K) The mesoderm‐specific RNAi‐based reduction of Dg and Dys (dsDys C‐term and dsDg30A/24B‐Gal4) at 20 days after eclosion, but not at 3 days after eclosion (I) show obvious IFM muscle pathology: the loss of fiber density and vacuolization (asterisks).

To determine whether this age‐dependent muscle degeneration phenotype is due to a requirement of Dg–Dys complex in muscle tissue, we used a mesoderm specific 24B‐Gal4‐driver to express the Dg and Dys RNAi constructs. Severe muscle degeneration phenotypes, accompanied with extensive vacuolization of muscle tissue and muscle fiber loss were observed when the Dg and Dys RNAi were directed in the mesoderm (Figure 4H–K; Table I). We further showed that, similar to what was observed in ubiquitous Dg and Dys RNAi animals (dsDg and dsDys/P‐tub Gal4), in muscle‐specific dsDg and dsDys/24B‐Gal4 mutants the muscle deterioration process has an age‐dependent character (Figure 4H–K; Table I). Taken together, these results suggest that, similar to human, Dg and Dys are required for muscle maintenance throughout the lifetime of Drosophila.

Dg and Dys are required for proper photoreceptor axon path‐finding

Brain‐selective deletion of Dg in mice is sufficient to cause congenital muscular dystrophy‐like brain malformations, including disarray of cerebral cortical layering and aberrant migration of granule neuronal cells (Michele et al, 2002; Moore et al, 2002; Qu and Smith, 2004). Within the cortex, however, it is not clear whether the Dg–Dys complex is required in neurons, glia, or both for proper neuronal migrations. To better understand the function of the Dg complex in the brain, we analyzed potential brain defects in the Drosophila Dg and Dys mutants.

Dg is expressed in the Drosophila adult eye, brain, and the developing larval brain and visual system, especially in optic lobes and photoreceptors (Figure 5B; Supplementary Figure 4D). In the optic lobe, Dg is present both on photoreceptor axons (in the optic stalk, lamina plexus and medulla neuropil) and the Repo‐expressing brain glial cells (Figure 5B). Dys shows similar expression patterns in the optic lobes.

To examine the role of Dg–Dys complex in the Drosophila brain, we analyzed frontal sections of adult heads from mutant Dys and Dg adult flies and observed abnormalities in the formation of retina: retinal photoreceptor cells were not elongated in Dg or Dys mutants (eyFLP; Dg323FRT 42D/FRT 42D l(2)cl‐R11 100%, Dys8‐2/DysKX43 88%, dsDg30A and dsDg33A/P‐tubGal4 92%, dsDysC‐term/P‐tubGal4 100%, control UAS GFP/P‐tubGal4 0%; Supplementary Figure 4), suggesting that the Dg–Dys complex is required in these photoreceptor sensory neurons.

The Drosophila compound eye consists of ∼800 ommatidia, each containing eight different photoreceptor sensory neurons, R cell subtypes that project axons into one of two optic ganglia layers in the brain during late larval development. R1–R6 axons innervate the most superficial layer, the lamina, generating a smooth lamina plexus, whereas R7 and R8 project axons through the lamina into the deeper medulla layer (Figure 5A and D) (Perez and Steller, 1996; Tessier‐Lavigne and Goodman, 1996; Clandinin and Zipursky, 2002; Ruan et al, 2002). The patterning of the R‐cell subtypes in eye discs and the extension of their axons to the optic lobes of the developing brains occur by late third instar larvae, while the elongation of the retinal cell body takes place at pupal stage (Izaddoost et al, 2002).

To determine at which stage Dg is required in photoreceptor neuron development, we induced eye‐specific mutant clones (eyFLP; Dg323FRT 42D/FRT 42D l(2)cl‐R11) and analyzed the developing neurons in late third instar larvae using a photoreceptor‐specific monoclonal antibody 24B10. The patterning of the Dg mutant ommatidia was normal, suggesting that Dg is not required for the determination and differentiation of the R‐cells. However, the axonal projections of these sensory neurons to the brain optic lobes were disturbed due to the lack of Dg, most of the axons migrate to the correct termination zone in lamina, but formed abnormal patches in the lamina plexus. Similar axonal problems were observed in Dys mutants. In the normal wild‐type brain the photoreceptor axons terminate in a stereotypic fashion producing a fan‐like structure in the lamina plexus (Figure 5A, B and D). However, in 85% of Dg loss‐of‐function (eyFLP; Dg323FRT 42D/FRT 42D l(2)cl‐R11) and 67–75% of Dys loss‐of‐function (Dys8‐2/DfKX43 and Dyse6/Dys8‐2) mutant third‐instar larvae optic lobes the lamina plexus is irregular; photoreceptor axons stop irregularly making gaps in the normal termination zone of the lamina plexus, deviate from the path and bundle aberrantly (Figure 5C, E and G; Table I).

Importantly, Dg and Dys proteins interact in controlling the photoreceptor axon path‐finding since simultaneous reduction of the level of both genes (Dg323/+; DysKX43/+ and Dg323/+; Dys8‐2/+) results in a high percentage of the axon projection phenotypes while reduction of each gene independently (Dg323/+, DysKX43/+ or Dys8‐2/+) does not (Figure 5F and C).

Dg and Dys are required both in neurons and glia for regular lamina plexus formation

We first showed that in the majority of Dg and Dys RNAi mutants driven by P‐tub‐Gal4, the photoreceptor axons exhibited targeting phenotypes similar to Dg clonal phenotypes, they bundled and/or terminated irregularly in the normal termination zone of the lamina plexus. When these Dys and Dg RNAi constructs were expressed in eye disks, photoreceptor axons similarly terminated irregularly in the lamina region of the brain and formed uneven lamina neuropil with gaps and abnormally densely packed regions (Figure 6A and B; Table I, dsDg and dsDys/GMR‐Gal4 74 and 61%). When Dys and Dg RNAi constructs were expressed in all glial cells, including eye disk and lamina glia, but not neurons, axons of the photoreceptor sensory neurons also showed bundling and irregular termination (Figure 6A and C; Table I, dsDg and dsDys/repo‐Gal4 71 and 76%). To test whether the obtained axon path‐finding phenotype is specific to DGC function in neurons and glia, we knocked‐down Dg and Dys in mesodermal tissue and observed no effect on the axon termination process above control samples (Figure 6A and D; dsDg and dsDys/24B‐Gal4). To determine the potential effect of DGC mutations on the development of laminal glial cells, we stained the third‐instar optic lobe using a monoclonal antibody that recognizes the glial‐specific nuclear protein Repo (Perez and Steller, 1996; Poeck et al, 2001). In wild type (Figure 6E and G), differentiating glial cells migrate into the lamina forming two clearly separated layers of glial cells (i.e., epithelial and marginal glia), which in turn present a stop signal for terminating R1–R6 growth cones in the lamina (Poeck et al, 2001). In Dg and Dys mutants, although glial cells migrated correctly into the lamina, they appeared less organized lacking the clear separation of epithelial and marginal glial layers (Figure 6F and H). We also used the MARCM technique in order to generate marked photoreceptor neurons and/or glial cells mutant for Dg323(elav‐Gal4 hsFLP;FRT42B tubGal80/FRT42BDg323;UAS GFP act<CD2<Gal4). The termination zone observed for mutant photoreceptors was irregular; clumping of axons at the lamina and lamina breaks were associated with the presence of Dg mutant glial cells as well as mutant photoreceptor axons (Supplementary Figure 5A and B). In contrast to the wild‐type regular axon/glia/axon pattern, in Dg mutant lamina the gaps were occupied by mislocalized glial cells (Supplementary Figure 5B). These data suggest that Dg acts autonomously and non‐autonomously for correct axon path‐finding; Dg–Dys complex is required both in neurons and in glial cells for proper neuron axonal growth and targeting.

Dg–Dys complex is required in both neurons and glial cells for proper axon path‐finding in Drosophila. (A) A bar graph represents the frequency of irregular and uneven lamina layer in Dg and Dys mutants. Analysis of axon path‐finding phenotypes using photoreceptor (GMR‐Gal4) and glia specific (repo‐Gal4) drivers suggests that Dg–Dys complex is required in neurons and glial cells; lack of Dg–Dys complex in either cell type results in axonal mistargeting. Knocking out of DGC in the mesoderm (24B‐Gal4) does not affect axon path‐finding over control levels. (B) A representative image of the majority of preparations showing the clumping and uneven lamina plexus phenotype (indicated by arrows) in photoreceptor specific Dg and Dys mutants (dsDg and dsDys/GMR‐Gal4). (C) Similar phenotypes observed when Dys and Dg RNAi constructs were expressed in all glial cells, including eye disk and lamina glia, but not neurons (dsDg and dsDys/Repo‐Gal4). (D) Axons of photoreceptor sensory neurons in dsDg and dsDys crossed to a muscle‐specific 24B‐Gal4 driver showed regular lamina layer, indistinguishable from control (Figure 5D). (E, F) In wild‐type (E) glial cells (marked with Repo) migrate from progenitor regions into the lamina where they are organized into two layers, the epithelial (e.g.) and marginal glia (m.g.), presenting a stop signal for the termination of R1–R6 growth cones at the lamina plexus (brackets). (F) In DGC mutants, although glial cells migrated correctly into the lamina, they appeared less organized. (G) In wild‐type Drosophila brain the termination zone is organized stereotypically: each axon terminates between glial cells resulting in a regular axon/glia/axon pattern. (H) In Dg or Dys mutants the termination zone is disorganized: glial cells are irregularly positioned and photoreceptor axons bundle causing gaps and densely packed regions in the lamina.

As discussed, several congenital muscular dystrophies exhibit neuronal migration disorders (Michele et al, 2002; Moore et al, 2002). The mediations of axon path‐finding and neuronal migration require similar processes including supportive glial cells (Bloch‐Gallego et al, 2005). In the vertebrate brain, Dg is required for granule neuron migration (Michele et al, 2002; Moore et al, 2002; Qu and Smith, 2004). It will be interesting to see in the future if similar to Drosophila axon path‐finding, the Dg–Dys complex in vertebrates acts both in neurons and glial cells for this process. Indeed, Dg function has been demonstrated in a support cell type in peripheral nervous system, Schwann cells for neuronal connectivity (Saito et al, 2003).

Discussion

The fly genome contains many highly conserved orthologues to human disease genes (Reiter et al, 2001; Bier, 2005), including neurological, cardiovascular, endocrine and metabolic disease‐genes. Among these, nearly all components of the Dg–Dys complex, which is involved in muscular dystrophies, are present in flies (Deng and Ruohola‐Baker, 2000; Greener and Roberts, 2000; Deng et al, 2003). We now show that Dys and Dg interact genetically and biochemically and are required in the same cell types in Drosophila. A fluorescence polarization assay revealed that the Dg–Dys binding interface is highly conserved in humans and Drosophila (Figure 2). Both proteins are required for oocyte cellular polarity and interact in this process (Figure 1). Futhermore, mutants of both Dg and Dys genes show symptoms observed in muscular dystrophy. Reduction of Dg and Dys proteins results in age‐dependent mobility defects (Figure 3). Eliminating Dg and Dys specifically in mesoderm derived tissues reveals that these proteins are required for muscle maintenance in adult flies: age‐dependent muscle degeneration was observed in mutant tissues (Figure 4). Dg–Dys complex is also required for neuron path‐finding and has both cell autonomous and non‐cell autonomous functions for this process (Figures 5 and 6). Further, we have now shown that in neuronal path‐finding process Dg interacts with InR and an SH2/SH3‐domain adapter molecule Nck/Dock (Figure 7).

Drosophila as a muscular dystrophy model

Animal models have been used efficiently in muscular dystrophy studies. Some of the models are naturally occurring mutations (mdx‐mouse, muscular dystrophy dog, cat and hamster), others have been generated by gene targeting (Watchko et al, 2002). However, the regulation and the control of Dg–Dys complex are not understood, and no successful therapeutics exist yet for muscular dystrophies (however, systemic delivery‐studies using adeno‐associated viral vectors show promise (Gregorevic et al, 2004)). Studies in new model organisms with easy‐to‐manipulate genetics might reveal the mode of regulation of the complex by identifying key regulatory components through suppressor screens. In addition, careful functional analysis of the complex in different cell types in model organisms might result in a unifying theme that will reveal its molecular mechanism of function. Such recently developed models for muscular dystrophy exist in C. elegans and zebrafish (Gieseler et al, 2000; Parsons et al, 2002; Bassett and Currie, 2003). In C. elegans Dys mutant, the transporter snf‐6 that normally participates in eliminating acetylcholine from the cholinergic synapses, is not properly localized, resulting in an increased acetylcholine concentration at the neuromuscular junction and muscle wasting (Kim et al, 2004). The function of Dys in neuromuscular junctions has also been recently addressed in Drosophila (van der Plas et al, 2006). These results bring up the possibility that muscular dystrophies in humans might also at least partly be attributed to the altered kinetics of acetylcholine transmission through neuromuscular junctions.

We have now shown that Drosophila melanogaster acts as a remarkably good model for age‐dependent progression of muscular dystrophy. Dg and Dys reduction in Drosophila show age‐dependent muscle degeneration and lack of climbing ability. It is tempting to speculate that the common denominator between different defects observed in Dg–Dys mutants in Drosophila and C. elegans is defective cellular polarity. The defects observed in C. elegans could be due to a defect in polarization of a cell, which will generate a neuromuscular junction that leads to miss‐targeted snf‐6. Similarly, we have shown that Drosophila Dg–Dys complex is required for cellular polarity in the oocyte. In addition, neural defects observed are plausibly due to polarity defects in the growing axon.

Dg–Dys complex in axon path‐finding

Similar to neuronal defects observed in human muscular dystrophy patients, neuronal defects were also found in Drosophila Dg and Dys mutant brains. In vertebrate brains, Dg affects neuronal migration (Montanaro and Carbonetto, 2003; Qu and Smith, 2004) possibly through interaction of neurons with their glial guides. The neuronal migration and process outgrowth have been shown to require supportive input from glial cells and involve the formation of adhesion junctions along the length of the soma. Also, the outgrowth of the leading process involves rapid extension and contraction over the length of the glial fiber (Rivas and Hatten, 1995; Shaham, 2005). Disruption of the cytoskeletal organization within the neuron, either of actin filaments (Rivas and Hatten, 1995) or microtubule interactions (Vallee et al, 2000), has been shown to inhibit glial‐mediated neuronal migration. The glial function in this process is less well studied.

Drosophila photoreceptor path‐finding provides an excellent system for genetic dissection of neuronal outgrowth and target recognition (Dickson, 2002). During the formation of the nervous system, newly born neurons send out axons to find their targets. Each axon is led by a growth cone that responds to extracellular axon guidance cues and chooses between different extracellular substrates upon which to migrate. Recent work has also identified a variety of intracellular signaling pathways by which these cues induce cytoskeletal rearrangements (Guan et al, 1996; Rao, 2005), but the proteins connecting signals from cell surface receptors to actin cytoskeleton have not been clearly determined. Dg is a good candidate for linking receptor signaling to the remodeling of the actin cytoskeleton and thereby polarizing the growth cone. We have now shown that perturbation of Dg–Dys complex causes phenotypes that resemble Nck/Dock‐Pak‐Trio axon path‐finding phenotypes (Figure 5) (Rao, 2005), suggesting that Dg may be one of the key players in Nck/Dock signaling pathway for axon guidance and target recognition in Drosophila.

Interestingly, Insulin receptor‐tyrosine kinase (InR) mutants also show similar phenotypes to those of Nck/Dock signaling in photoreceptor axon path‐finding and these two proteins show genetic and biochemical interactions (Song et al, 2003). These data have led to speculations of mammalian InR acting in conjunction with Nck/Dock pathway in learning, memory and eating behavior (Dickson, 2003; Song et al, 2003). Our data now add Dg–Dys complex to this pathway; similar to what is seen in the case of Dg and Dys photoreceptor mutants, InR mutants show no obvious defects in patterning of the photoreceptors. However, the guidance of photoreceptor cell axons from the retina to the brain is aberrant (Song et al, 2003; Figures 5 and 6). Furthermore, genetic and biochemical evidence suggests that InR function in axon guidance involves the Dock‐Pak pathway rather than the PI3K‐Akt/PKB pathway. Independently, biochemical interaction between Nck/Dock and Dg has been reported (Sotgia et al, 2001) supporting the hypothesis that InR, Dg and Nck/Dock interaction regulates Dg–Dys complex. Furthermore, we have now shown that Dg interacts genetically with InR and Dock in photoreceptor axon path‐finding. Since Dys interacts with Dg but not with InR and Dock, it is tempting to speculate that Dg can selectively interact with either Dys or InR and Dock (Figure 7). One possibility is that the tyrosine kinase activity of InR could regulate the Dg–Dys interaction by tyrosine phosphorylation in the Dg–Dys binding interphase (Figure 2). This tyrosine phosphorylation could prohibit the Dg–Dys interaction and thereby result in rearrangements in the actin cytoskeleton. Alternatively, other components observed in Dg–Dys complex might be involved in this regulation (Zhan et al, 2005). However, it is also possible that potential polarity defects in the Dg mutant axons result in defective InR membrane localization. Interestingly, in another cell type, the Drosophila oocyte, InR, Dg and Dys also show similar phenotypes (Deng et al, 2003; LaFever and Drummond‐Barbosa, 2005; Figure 1). In addition, insulin‐like growth factors (IGF) and InR are important in maintaining muscle mass in vertebrates (Singleton and Feldman, 2001). Further connection of InR to Dg–Dys complex comes from experiments showing that muscle specific expression of IGF counters muscle decline in mdx‐mice (Barton et al, 2002; Shavlakadze et al, 2004; Dobrowolny et al, 2005). The work presented in this study is the first demonstration of genetic interaction between Dg and InR. Future biochemical studies should unravel the molecular mechanism of this interaction.

Furthermore, we have now shown that Dg–Dys complex is required both in neural and in targeting glial cells for correct neuronal axon path‐finding in Drosophila brain. These data reveal that Dg–Dys complex also has a non‐cell autonomous effect on axon path‐finding and suggest that Dg–Dys‐controlled ECM both from neuron and glial cells regulate neuronal axon path‐finding. Further experiments are required to reveal whether long‐range Laminin fibers are involved in this process, as has been shown in epithelial planar polarity (Bateman et al, 2001; Deng et al, 2003), or whether glial processes are observed in close proximity to the neural growth cone (Georges‐Labouesse et al, 1998). Interestingly, similar phenotypes are observed with Integrin mutants (Tanaka and Sabry, 1995; Campos, 2005; Curtin et al, 2005), suggesting that, as in planar polarity (Bateman et al, 2001; Deng et al, 2003), Integrin and Dg–Dys complex might act in concert to regulate the process of ECM organization that will regulate the cytoskeleton of the cells involved.

Taken together, the phenotypes caused by Drosophila Dg and Dys mutations are remarkably similar to phenotypes observed in human muscular dystrophy patients, and therefore suggest that functional dissection of Dg–Dys complex in Drosophila should provide new insights into the origin and potential treatment of these fatal neuromuscular diseases. As a proof of principle, using Drosophila as a model we have now identified InR as a signaling pathway that genetically interacts with Dg. Future studies are directed to unravel the molecular mechanism of Dg and InR–Dock interactions in invertebrates as well as vertebrates.

Supplementary data

Supplementary Information

Acknowledgements

We thank Drs Leslie Pick and Paul Garrity for reagents and advice, Alex Whitworth for help with mobility assay, Glenda Froelick for help with histological sections, Volodymyr Shcherbatyy for injections to produce the transgenic lines, Christian Walker‐Richards and Luis Tulloch for generating Dys8‐2 deletion line, Merle Gilbert for help with initial experiments, Greg Martin for confocal microscopy instructions, Junlin Qi for help with qRT–PCR, Arul Subramanian and Amir Sapir for helping in the preparation of the RNAi flies, Michael Eck and Florence Poy for the human Dystrophin clone, Martina Schneider and Ellen Ward for help with embryonic analysis, and Mario Pantoja for critical reading. This work was supported by an AHA fellowship for HRS, CRDF for ASY, HRS and HR‐B, CIHR for VDS, MDA, AFM grants and The Israel Science Foundation for UN and DY, MDA for HR‐B and by the grants from the National Institute of Health for DB and HR‐B.